Name: in vivo interrogation of central nervous system translatome by polyribosome fractionation
Uploaded: 2014-04-30T13:00:00
Duration: 9 min 13 s
Description: Watch this Scientific Journal Video about In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation at JoVE.com

This work was supported by the German Federal Ministry of Education and Research (BMBF), the Systems Biology of Signaling in Cancer (Helmholtz Alliance on Systems Biology) and the German Cancer Research Center (DKFZ).

All animal experiments were performed in accordance with the institutional guidelines of the DKFZ. 
Caution: In order to prevent RNAse contamination of samples, take basic precautions for avoiding RNAse contamination and prepare all buffers with DEPC-treated water.
1. Preparation of Sucrose Gradients
<ol>
	<li>Prepare 17.5, 25.6, 33.8, 41.9 and 50% sucrose solutions (see Table 1). Start preparing gradients by slowly adding 2 ml of 50% sucrose solution to the bottom of a polyallomer ultracentrifuge tube. Freeze the layer by placing tubes for 20 min at -80 &#176;C.</li>
	<li>Add the subsequent layers at decreasing sucrose percentages with freezing steps in between, ending up to 17.5% sucrose. Keep gradients on ice during addition of sucrose solutions in order to prevent thawing of underlying layers.</li>
	<li>Prepare sucrose gradients freshly one day before use and keep at 4 &#176;C. Alternatively, prepare gradients in advance, store at -80 &#176;C and thaw at 4 &#176;C the night before the experiment.</li>
</ol>
2. Tissue Preparation (Spinal Cord and/or Brain)
<ol>
	<li>Anesthetize mouse by an overdose of Xylazin and Ketamin in NaCl and transcardially perfuse with&#160;20 ml of Hank&#39;s balanced salts solution (HBSS) containing 200 &#181;g/ml CHX which immobilizes ribosomes on associated transcripts. Confirm proper anesthetization by non-responsiveness to toe pinches.</li>
	<li>Extract tissue rapidly. Open the cranium dorsally and extract the brain. Perform laminectomy of the entire thoracic and lumbar vertebral column and extract the spinal cord. Use whole brain (~400 mg) or a 4 cm piece of spinal cord (~90 mg), respectively.</li>
	<li>Transfer tissue into ice cold homogenization buffer A (see Table 1) (spinal cord: 1 ml; brain: 4 ml) and dice into small pieces with a clean scalpel to allow increased uptake of cycloheximide. Perform all further steps on ice.</li>
	<li>Incubate for 15 min and homogenize tissue mechanically using a Dounce homogenizer. Carefully controlled homogenization is necessary to ensure that nuclei remain intact. Treat all samples exactly the same way to ensure comparability. Note: for brain tissue:&#160;Disrupt tissue by 5 strokes with a tightly fitting pestle. For spinal cord tissue: disrupt by 5 strokes with a loosely fitting pestle, followed by 5 further strokes with a tightly fitting pestle.</li>
	<li>Take aliquots (400 &#181;l for brain and 200 &#181;l for spinal cord), flash freeze and store at -80 &#176;C for total RNA isolation.</li>
	<li>Remove nuclei and undisrupted cell and tissue fragments by centrifugation at 500 x g for 10 min at 4 &#176;C. Low speed centrifugation prevents loss of ribosomes.</li>
	<li>Lyse membrane fragments in the nuclei free supernatant for 30 min by adding NP-40 and sodium deoxycholate detergents (each to a final concentration of 1%).</li>
</ol>
3. Myelin Flotation
<ol>
	<li>This step removes fatty components from the sample which will otherwise mask the signal in the polyribosome profile. First, pre-chill ultracentrifuge buckets and rotor at 4 &#176;C and prepare 2 M, 1.1 M and 0.9 M sucrose solutions (see Table 1).</li>
	<li>Mix lysate with an 1.22&#160;volumes of 2 M sucrose solution and transfer into a polyethylene ultracentrifuge tube. Fill up all tubes to equal volume of 10 ml with 1.1 M sucrose solution, thereafter overlay it carefully with 0.9 M sucrose solution.</li>
	<li>Place the ultracentrifuge tubes into pre-chilled ultracentrifuge buckets. Centrifuge for 3 hr at 100,000 x g at 4 &#176;C. During this process ribosomes become deposited in the pellet whereas fatty components float up and remain in the supernatant.</li>
</ol>
4. Sucrose Gradient Fractionation
<ol>
	<li>Remove supernatant and dissolve pellet in homogenization buffer B (see Table 1). Measure the absorbance at 260 nm for each sample using a Nanodrop or equivalent apparatus and normalize amounts to load according to the absorbance value.</li>
	<li>Place sucrose gradients (preparation is described in a previous section) into the pre-chilled ultracentrifuge buckets. Layer samples carefully on top of the gradient. Add one blank sucrose gradient as technical control.</li>
	<li>Adjust the weight of each bucket with homogenization buffer. Centrifuge samples for 1.5 hr at 285,000 x g at 4 &#176;C.</li>
	<li>Start preparing the Isco fractionator 30 min before the end of centrifugation. If another model of fractionator is used, follow the manufacturer&#39;s protocol.
	<ol>
		<li>Set the appropriate sensitivity (0.2 AUFS for brain or 0.05 AUFS for spinal cord) for the UV lamp and switch it on for warming up. Assemble the tube piercer, connect it via the rolling pump to 60% sucrose solution through the tube piercer.</li>
		<li>Test the setup by pumping sucrose (flow rate: 1 ml/min), in particular make sure there is no leaks in the tubings which will introduce bubbles into the gradient.</li>
		<li>Blank background absorption by manually pumping gradient buffer into the UV detector and correcting the baseline.</li>
	</ol>
	</li>
	<li>Carefully remove the ultracentrifuge buckets containing the sucrose gradients with sedimented samples from the rotor and place them on ice. Avoid any bumping of gradients to prevent loss of resolution.</li>
	<li>Run gradients on the fractionator. Run empty gradients first to assess background absorption and ensure proper technical setup.
	<ol>
		<li>Attach gradient to the UV detector and pierce the bottom of the tube with the tube piercer. Start the pumping of 60% sucrose, which will displace the gradient with sedimented samples upwards through the UV detector and into the drop dispenser.</li>
		<li>The absorbance at 260 nm is documented to generate the absorption profile for the sample, with peaks indicating the sedimentation of mRNAs associated with ribosomal subunits, monosomes, and subsequently increasing number of ribosomes.</li>
	</ol>
	</li>
	<li>Collect samples into 20 fractions via the drop dispenser (600 &#181;l each, in 2 ml tubes).</li>
</ol>
5. RNA Isolation from Individual Fractions
<ol>
	<li>Add 10% SDS to a final concentration of 1% to individual fractions and mix well in order to unfold proteins and dissociate ribosomes. At this point samples can be frozen at -80 &#176;C. Depending on the question to be addressed, fractions can be pooled according to the absorbance profile.</li>
	<li>Isolate RNA with acidic phenol/chloroform extraction, which also removes contaminating traces of DNA.
	<ol>
		<li>Add one volume of acidic phenol/chloroform (prewarmed to RT) to each sample, heat for 10 min at 65 &#176;C and release pressure under the fume hood afterwards.</li>
		<li>Centrifuge samples for 20 min at 17,000 x g at RT. Carefully transfer aqueous phase to a new 1.5 ml tube. Be aware of phase inversion in denser sucrose fractions, where the aqueous phase might be at the bottom after centrifugation.</li>
		<li>Add one volume of isopropanol, 1/9 volume sodium acetate (pH 5.2) and 1&#160;&#181;l GlycoBlue to each sample in order to precipitate RNA. Keep samples at least 1 hr at -80 &#176;C.</li>
		<li>Centrifuge for 30 min at 17,000 x g and 4 &#176;C. GlycoBlue provides visualization of the pellet. Remove supernatant, wash pellet once with ice cold 80% ethanol and air dry pellet. Dissolve pellet in RNAse free water.</li>
	</ol>
	</li>
	<li>Quantify amount of RNA by Nanodrop and analyze RNA integrity by a Bioanalyzer chip. RNA which is obtained by the presented protocol is suitable for all state of the art high throughput assays including microarray&#160;and deep sequencing.</li>
</ol>

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M., Foissac, S., Guig&#243;, R., M&#233;ndez, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+combinatorial+code+for+CPE-mediated+translational+control.">A combinatorial code for CPE-mediated translational control.</a> Cell. 132 (3), 434-448 (2008).</li><li>Ingolia, N. T., Ghaemmaghami, S., Newman, J. R. S., Weissman, J. 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No conflict of interest declared.

Although polyribosome fractionation is not a novel technique, it remains a particularly challenging one. Based on the input material, substantial optimization can be necessary. This is especially the case for in vivo CNS samples, where the amount of material is often a limitation and fatty tissue components hinder isolation of translating mRNAs. Most published fractionation protocols deal with yeast or mammalian cell lines, and there are established protocols for the brain12,13,14. In contrast, there are barely any publications describing fractionation of the spinal cords, and previous protocols require spinal cord&#160;from a large number of animals to be pooled15. For these reasons, several essential modifications were made to adapt the fractionation protocol for CNS tissues, including single mouse spinal cord. Ribosome immobilization with cycloheximide is performed during animal perfusion to avoid dissociation of ribosomes during the long process of tissue extraction. Subsequently, polysomal RNAs are extracted in a specific manner to maximize polyribosome yield. First, controlled homogenization of the tissue by douncing, keeps the nucleus intact and prevents DNA contamination. The nucleus is then removed reliably by centrifugation. The combination of the detergents NP-40 and sodium deoxycholate ensures lysis of the endoplasmic reticulum and release of its membrane associated ribosomes. In addition, UV-absorbing components within the lipid rich myelin obscure the polyribosome profiles. Myelin flotation is therefore necessary, where dense compounds such as polyribosomes are pelleted and lighter compounds such as myelin float and are removed16. Polyribosomes are then sedimented over a 17.5% to 50% sucrose gradient. Instead of manually layering and freezing each sucrose layer, gradients can also be prepared using a gradient mixer. Extraction of RNA from fractions using acidic phenol/chloroform allows DNA to be removed with minimal RNA loss. However, phase inversion may occur at dense sucrose fractions (above fraction 16).
This protocol provides advantages over other conventional methods that address the level of translation. For example, both the measurement of phosphorylated S6 (a prominent translation marker) as well as labeling of nascent chains with radioisotopes give information on global translation levels but reveal little on what specifically is being translated. Polyribosome fractionation, on the other hand, allows not only assessment of global translation, but also of the identity of translating mRNAs and the related regulatory proteins. Quantitative real time PCR can be performed on isolated RNAs for a quick read out of selected mRNAs, and microarrays and next generation RNA sequencing can be performed for genome wide studies. The optimizations presented here allow the technique to be used with minimal amounts of CNS tissues, therefore diminishing the number of animals needed and improving overall experimental quality by reducing tissue processing time. On the other hand, this technique cannot distinguish stalled ribosomes from translating ones. Transcripts carrying stalled ribosomes will sediment in the heavy fractions, and this should be kept in mind when interpreting the data.
There are recent reported techniques, namely RiboTaq and TRAP, in which ribosomes are labeled with epitope tags or reporters respectively in a cell type specific manner and associated mRNAs isolated by immunoprecipitation17,18. This technique can be coupled to polyribosome fractionation to offer high resolution read out for specific cell populations. Ribosome foot-printing is another novel technique that involves nuclease digestion to generate small fragments, or &#8220;footprints&#8221;, of mRNAs that are protected by ribosomes. Libraries are then generated from these footprints and sequenced. This method provides a codon-specific whole genome analysis on translation and is able to identify stalled ribosomes, non-AUG start codons and small upstream open reading frames, which cannot be achieved by polyribosome fractionation19. However, polyribosome fractionation can be coupled to ribosome profiling for isolation of digested fragments containing single ribosomes, thus enriching for the molecular species needed for library preparation in which high sample purity is often required. Taken together, polyribosome fractionation is a flexible method with lasting significance and can be coupled to various downstream applications including genome wide assays like next-generation sequencing.

Gene expression is determined by the combined action of transcription, translation and stability of mRNA and proteins, with translation bearing the most predominant effect1. It is now evident that each of these steps is highly regulated. Micro-RNAs, formation of RNA granules, alternative and cytoplasmic polyadenylation are some examples of post-transcriptional regulation of gene expression2,3. Each of these mechanisms uncouples transcription from translation and influences the proteome in the biological system of interest. Thus, unsurprisingly, mRNA levels alone are an imperfect readout of protein levels4. Quantitative proteomics provides the most direct assessment of gene expression, however despite recent advances, there are still considerable limitations on sensitivity and protein sequence resolution. Therefore addressing the translatome, the repertoire of translating mRNAs, offers an excellent compromise between studying the transcriptome and proteome. It is more accurate than transcriptomics in assessing final gene expression, and provides both higher coverage and sequence resolution than proteomics.
In mammalian systems, the majority of translation events begin via cap-dependent initiation. A group of eukaryotic initiation factors together with the 40S small ribosomal subunit assemble at the 5&#39; cap of an mRNA. The complex then scans the mRNA and upon reaching the start codon AUG, recruits the 60S large ribosomal subunit to form a complete 80S ribosome. The elongation stage proceeds with the ribosome moving along the mRNA, with elongation factors assisting the incorporation of amino acids from loaded tRNAs to the nascent peptide chain. Multiple ribosomes can proceed along a single mRNA simultaneously and the number of associated ribosomes has been shown to correlate with the rate of protein synthesis5,6. This makes ribosomal loading a reliable indication for translation and allows separation of actively translating mRNAs based on sedimentation rate. In addition to quantification of translating mRNAs, sequence information can be obtained to identify motifs involved in translation regulation. Also RNA binding proteins and other translation factors can be isolated from different fractions, facilitating the study and discovery of related regulatory proteins.
In the nervous system, translational control has been linked to processes such as mRNA storage, transport and local protein synthesis. Growth cones have been shown to harbor a specific localized pool of mRNAs distinct from the rest of the axon7. In addition, axons possess the ability to locally synthesize proteins8,9. As a result, local control of translation has become a crucial topic of study in neurobiology. The potential of polyribosome fractionation to address this has been illustrated in several studies, in which the technique was used to investigate axon guidance in spinal cord development, and demonstrated the activity-dependent translation of BDNF in the brain10,11.

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		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Hank&#8217;s balanced salts solution</td>
			<td style="width:147px;">Gibco&#160;</td>
			<td style="width:119px;">14170-138</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tube, Thinwall, Polyallomer, 14 mL, 14 x 95mm</td>
			<td>Beckman Coulter</td>
			<td>331374</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Diethylpyrocarbonate&#160;</td>
			<td>Sigma</td>
			<td>D5758</td>
			<td>caution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cycloheximide</td>
			<td>Sigma</td>
			<td>C7698</td>
			<td>danger</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dithiothreitol</td>
			<td>Sigma</td>
			<td>43815</td>
			<td>warning</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">cOmplete Protease Inhibitor Cocktail Tablets</td>
			<td>Roche</td>
			<td>11697498001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Phenylmethanesulfonyl fluoride solution</td>
			<td>Sigma</td>
			<td>93482</td>
			<td>danger</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">RNasin Plus RNase Inhibitor</td>
			<td>Promega</td>
			<td>N2611</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Nonidet P-40</td>
			<td>Roche</td>
			<td>11332473001</td>
			<td>danger</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium deoxycholate</td>
			<td>Sigma</td>
			<td>D6750</td>
			<td>warning</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Acid-Phenol:Chloroform, pH 4.5 (with IAA, 125:24:1)</td>
			<td>Ambion</td>
			<td>AM9722</td>
			<td>danger</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">GlycoBlue</td>
			<td>Invitrogen</td>
			<td>AM9516</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">Equipment</td>
			<td style="width:147px;"></td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dounce Tissue Grinder, 1mL/7ml</td>
			<td>Wheaton</td>
			<td>357538/357542</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Nanodrop 2000</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">SW 40 Ti Rotor, Swinging Bucket, Titanium, 6 x 14 mL, 40,000 rpm, 285,000 x&#160;g</td>
			<td>Beckman Coulter</td>
			<td>331302</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Density Gradient Fractionator</td>
			<td>Teledyne Isco</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:380px;">2100 Bioanalyzer</td>
			<td style="width:147px;">Agilent Technologies</td>
			<td style="width:119px;"></td>
			<td></td>
		</tr>
	</tbody>
</table>

in vivo interrogation of central nervous system translatome by polyribosome fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are tightly regulated, affecting the final dynamics of protein abundance. Various regulatory mechanisms exist at the translation step, rendering mRNA levels alone an unreliable indicator of gene expression. In addition, local regulation of mRNA translation has been particularly implicated in neuronal functions, shifting &#39;translatomics&#39; to the focus of attention in neurobiology. The presented method can be used to bridge transcriptomics and proteomics.
Here we describe essential modifications to the technique of polyribosome fractionation, which interrogates the translatome based on the association of actively translated mRNAs to multiple ribosomes and their differential sedimentation in sucrose gradients. Traditionally, working with in vivo samples, particularly of the central nervous system (CNS), has proven challenging due to the restricted amounts of material and the presence of fatty tissue components. In order to address this, the described protocol is specifically optimized for use with minimal amount of CNS material, as demonstrated by the use of single mouse spinal cord and brain. Briefly, CNS tissues are extracted and translating ribosomes are immobilized on mRNAs with cycloheximide. Myelin flotation is then performed to remove lipid rich components. Fractionation is performed on a sucrose gradient where mRNAs are separated according to their ribosomal loading. Isolated fractions are suitable for a range of downstream assays, including new genome wide assay technologies.

Figure 2 shows representative polyribosome profiles after fractionation. The profiles of brain and spinal cord show characteristic absorption curves as described before for cell lines. mRNAs bound to small ribosomal subunits (40S) sediment at lighter fractions and appear first as a peak on the profile, followed by the large ribosomal subunit (60S) and monosome (80S) bound mRNAs. mRNAs bound to multiple ribosomes sediment at heavier fractions, with the later peaks indicating increasing number of bound ribosomes. Global translation can be assessed from the profile. As an example, brain tissue has a higher polyribosome to monosome ratio than spinal cord tissue, indicating more active translation.
RNA extracted from individual fractions were assessed by Bioanalyzer, showing distribution of 18S and 28S rRNA. 18S rRNA appears earlier in the profile, in accordance with the small ribosomal subunits sedimenting in lighter sucrose fractions. Typical yields of total RNA are 10-20 &#181;g for brain and 2-4 &#181;g for spinal cord. RNA yields from individual fractions are up to 4 &#181;g and 0.8 &#181;g for brain and spinal cord respectively, depending on the fraction. A blank sucrose gradient already shows some background absorption at 260 nm, due to the presence of DTT. This background can be subtracted during data analysis in order to normalize sample values. EDTA treatment collapsed the polyribosome peaks, demonstrating the sedimentation profile is due to translation.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51255/51255fig1highres.jpg" width="500" />
Figure 1.&#160;Workflow and potential applications of in vivo polyribosome fractionation. <a href="https://www.jove.com/files/ftp_upload/51255/51255fig1highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" fo:content-width="5in" fo:src="/files/ftp_upload/51255/51255fig2highres.jpg" src="/files/ftp_upload/51255/51255fig2.jpg" /> 
Figure 2.&#160;(A) Typical polyribosome profiles of brain and spinal cord, with assessment of extracted RNA by Bioanalyzer. (B) Distribution of Gapdh mRNA over fractions by qRT-PCR. (C)&#160;Polyribosome profiles of empty gradient and brain with and without EDTA treatment. <a href="https://www.jove.com/files/ftp_upload/51255/51255fig2highres.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="0" cellpadding="0" cellspacing="0" fo:keep-together.within-page="always">
	<tbody>
		<tr>
			<td>1.1</td>
			<td>2x gradient buffer</td>
			<td>30 mM Tris-HCl pH7.4, 30 mM MgCl2, 600 mM NaCl, 200 &#181;g/ml cycloheximide (CHX), 2mM dithiothreitol (DTT)</td>
			<td></td>
		</tr>
		<tr>
			<td>1.1</td>
			<td>sucrose solution 17.5%</td>
			<td>8.67 g sucrose, 25 ml 2x gradient buffer, up to 50 ml DEPC H20&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>1.1</td>
			<td>sucrose solution 25.6%</td>
			<td>12.8 g sucrose, 25 ml 2x gradient&#160; buffer, up to 50 ml DEPC H20&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>1.1</td>
			<td>sucrose solution 33.8%</td>
			<td>16.9 g sucrose, 25 ml 2x gradient&#160; buffer, up to 50 ml DEPC H20&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>1.1</td>
			<td>sucrose solution 41.9%</td>
			<td>20.95 g sucrose, 25 ml 2x gradient &#160;buffer, up to 50 ml DEPC H20&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>1.1</td>
			<td>sucrose solution 50%</td>
			<td>25 g sucrose, 25 ml 2x gradient buffer, up to 50 ml DEPC H20&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>2.3</td>
			<td>homogenization buffer A</td>
			<td>0.25 M sucrose, 50 mM Tris/HCl, pH7.4), 5 mM&#160; MgCl2, 25 mM KCl</td>
			<td>200 &#181;g/ml CHX, 1x Roche complete protease inhibitor, 1 mM DTT, 1mM phenylmethanesulfonylfluoride (PMSF), 100 U/ml RNAsin&#160;</td>
		</tr>
		<tr>
			<td>3.1</td>
			<td>2M sucrose solution</td>
			<td>68.4% sucrose, 50 mM Tris-HCl, pH7.4, 5 mM MgCl2, 25 mM KCl</td>
			<td>100 &#181;g/ml CHX, 1x Roche complete protease inhibitor, 1 mM DTT, 1 mM PMSF</td>
		</tr>
		<tr>
			<td>3.1</td>
			<td>1.1M sucrose solution</td>
			<td>38.5% sucrose, 50 mM Tris-HCl, pH7.4, 5 mM MgCl2, 25 mM KCl</td>
			<td>100 &#181;g/ml CHX, 1x Roche complete protease inhibitor, 1 mM DTT, 1 mM PMSF</td>
		</tr>
		<tr>
			<td>3.1</td>
			<td>0.9M sucrose solution</td>
			<td>30.8% sucrose, 50 mM Tris-HCl, pH7.4, 5 mM MgCl2, 25 mM KCl</td>
			<td>100 &#181;g/ml CHX, 1x Roche complete protease inhibitor, 1 mM DTT, 1 mM PMSF</td>
		</tr>
		<tr>
			<td>4.1</td>
			<td>homogenization buffer B</td>
			<td>0.25 M sucrose, 50 mM Tris/HCl, pH7.4, 5 mM&#160; MgCl2, 25mM KCl, 1% NP-40, 1% sodium deoxycholate&#160;</td>
			<td>200 &#181;g/ml CHX, 1x Roche complete protease inhibitor, 1mM DTT, 1 mM PMSF&#160;&#160;, 100 U/ml RNAsin&#160;</td>
		</tr>
	</tbody>
</table>
Table 1.&#160;List of buffers and solutions.

Watch this Scientific Journal Video about In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation at JoVE.com

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

This protocol illustrates essential modifications of polyribosome fractionation in order to study the translatome of in vivo CNS samples. It allows global assessment of translation and transcription regulation through the isolation and comparison of total RNA to ribosome bound RNA fractions.

In vivo Interrogation of Central Nervous System Translatome by Polyribosome Fractionation

Multiple processes are involved in gene expression including transcription, translation and stability of mRNAs and proteins. Each of these steps are ...

in-vivo-interrogation-central-nervous-system-translatome-polyribosome

Research

JoVE Journal

Neuroscience

Optic Nerve Transection: A Model of Adult Neuron Apoptosis in the Central Nervous System

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be accessed within the orbit of the eye and completely transected (axotomized), cutting the axons of the entire RGC population. Optic nerve 
transection is a reproducible model of apoptotic neuronal cell death in the adult CNS 1-4. This model is particularly attractive because the vitreous
 chamber of the eye acts as a capsule for drug delivery to the retina, permitting experimental manipulations via intraocular injections. The diffusion of chemicals 
through the vitreous fluid ensures that they act upon the entire RGC population. Moreover, RGCs can be selectively transfected by applying short interfering RNAs 
(siRNAs), plasmids, or viral vectors to the cut end of the optic nerve 5-7 or injecting vectors into their target, the superior colliculus 8. 
This allows researchers to study apoptotic mechanisms in the desired neuronal population without confounding effects on other bystander neurons or surrounding glia. 
An additional benefit is the ease and accuracy with which cell survival can be quantified after injury. The retina is a flat, layered tissue and RGCs are localized in 
the innermost layer, the ganglion cell layer. The survival of RGCs can be tracked over time by applying a fluorescent tracer (3% Fluorogold) to the cut end of the 
optic nerve at the time of axotomy, or by injecting the tracer into the superior colliculus (RGC target) one week prior to axotomy. The tracer is retrogradely transported, labeling 
the entire RGC population. Because the ganglion cell layer is a monolayer (one cell thick), RGC densities can be quantified in flat-mounted tissue, without the need 
for stereology. Optic nerve transection leads to the apoptotic death of 90% of injured RGCs within 14 days postaxotomy 9-11. RGC apoptosis has a 
characteristic time-course whereby cell death is delayed 3-4 days postaxotomy, after which the cells rapidly degenerate. This provides a time window for 
experimental manipulations directed against pathways involved in apoptosis.

Optic Nerve transection is a widely used model of adult CNS injury. Ninety percent of retinal ganglion cells (RGCs) whose axons are completely transected (axotomy) die within 14 days after axotomy. This model is easily amenable to experimental manipulations and highly reproducible.

Retinal ganglion cells (RGCs) are CNS neurons that output visual information from the retina to the brain, via the optic nerve. 
The optic nerve can be ...

Multimodal Imaging of Stem Cell Implantation in the Central Nervous System of Mice

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, both in preclinical and clinical studies. However, to date results regarding functional outcome and/or tissue regeneration following stem cell transplantation are quite diverse. Generally, a clinical benefit is observed without profound understanding of the underlying mechanism(s)1. Therefore, multiple efforts have led to the development of different molecular imaging modalities to monitor stem cell grafting with the ultimate aim to accurately evaluate survival, fate and physiology of grafted stem cells and/or their micro-environment. Changes observed in one or more parameters determined by molecular imaging might be related to the observed clinical effect. In this context, our studies focus on the combined use of bioluminescence imaging (BLI), magnetic resonance imaging (MRI) and histological analysis to evaluate stem cell grafting.
BLI is commonly used to non-invasively perform cell tracking and monitor cell survival in time following transplantation2-7, based on a biochemical reaction where cells expressing the Luciferase-reporter gene are able to emit light following interaction with its substrate (e.g. D-luciferin)8, 9. MRI on the other hand is a non-invasive technique which is clinically applicable10 and can be used to precisely locate cellular grafts with very high resolution11-15, although its sensitivity highly depends on the contrast generated after cell labeling with an MRI contrast agent. Finally, post-mortem histological analysis is the method of choice to validate research results obtained with non-invasive techniques with highest resolution and sensitivity. Moreover end-point histological analysis allows us to perform detailed phenotypic analysis of grafted cells and/or the surrounding tissue, based on the use of fluorescent reporter proteins and/or direct cell labeling with specific antibodies.
In summary, we here visually demonstrate the complementarities of BLI, MRI and histology to unravel different stem cell- and/or environment-associated characteristics following stem cell grafting in the CNS of mice. As an example, bone marrow-derived stromal cells, genetically engineered to express the enhanced Green Fluorescent Protein (eGFP) and firefly Luciferase (fLuc), and labeled with blue fluorescent micron-sized iron oxide particles (MPIOs), will be grafted in the CNS of immune-competent mice and outcome will be monitored by BLI, MRI and histology (Figure 1).

This article describes an optimized sequence of events for multimodal imaging of cellular grafts in rodent brain using: (i) in vivo bioluminescence and magnetic resonance imaging, and (ii) post mortem histological analysis. Combining these imaging modalities on a single animal allows cellular graft evaluation with high resolution, sensitivity and specificity.

During the past decade, stem cell transplantation has gained increasing interest as primary or secondary therapeutic modality for a variety of diseases, ...

Production of Lentiviral Vectors for Transducing Cells from the Central Nervous System

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing therapeutic approaches. Lentiviral vectors are attractive tools in transduction of neurons and other cell types in CNS as they transduce both dividing and non-dividing cells, support sustained expression of transgenes, and have relatively large packaging capacity and low toxicity 1-3. Lentiviral vectors have been successfully used in transducing many neural cell types in vitro 4-6 and in animals 7-10.
Great efforts have been made to develop lentiviral vectors with improved biosafety and efficiency for gene delivery. The current third generation replication-defective and self-inactivating (SIN) lentiviral vectors are depicted in Figure 1. The required elements for vector packaging are split into four plasmids. In the lentiviral transfer plasmid, the U3 region in the 5' long terminal repeat (LTR) is replaced with a strong promoter from another virus. This modification allows the transcription of the vector sequence independent of HIV-1 Tat protein that is normally required for HIV gene expression 11. The packaging signal (&Psi;) is essential for encapsidation and the Rev-responsive element (RRE) is required for producing high titer vectors. The central polypurine tract (cPPT) is important for nuclear import of the vector DNA, a feature required for transducing non-dividing cells 12. In the 3' LTR, the cis-regulatory sequences are completely removed from the U3 region. This deletion is copied to 5' LTR after reverse transcription, resulting in transcriptional inactivation of both LTRs. Plasmid pMDLg/pRRE contains HIV-1 gag/pol genes, which provide structural proteins and reverse transcriptase. pRSV-Rev encodes Rev which binds to the RRE for efficient RNA export from the nucleus. pCMV-G encodes the vesicular stomatitis virus glycoprotein (VSV-G) that replaces HIV-1 Env. VSV-G expands the tropism of the vectors and allows concentration via ultracentrifugation 13. All the genes encoding the accessory proteins, including Vif, Vpr, Vpu, and Nef are excluded in the packaging system. The production and manipulation of lentiviral vectors should be carried out according to NIH guidelines for research involving recombinant DNA (<a href="http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf" target="_blank">http://oba.od.nih.gov/oba/rac/Guidelines/NIH_Guidelines.pdf</a>). An approval from individual Institutional Biological and Chemical Safety Committee may be required before using lentiviral vectors. Lentiviral vectors are commonly produced by cotransfection of 293T cells with lentiviral transfer plasmid and the helper plasmids encoding the proteins required for vector packaging. Many lentiviral transfer plasmids and helper plasmids can be obtained from Addgene, a non-profit plasmid repository (<a href="http://www.addgene.org/" target="_blank">http://www.addgene.org/</a>). Some stable packaging cell lines have been developed, but these systems provide less flexibility and their packaging efficiency generally declines over time 14, 15. Commercially available transfection kits may support high efficiency of transfection 16, but they can be very expensive for large scale vector preparations. Calcium phosphate precipitation methods provide highly efficient transfection of 293T cells and thus provide a reliable and cost effective approach for lentiviral vector production.
In this protocol, we produce lentiviral vectors by cotransfection of 293T cells with four plasmids based on the calcium phosphate precipitation principle, followed by purification and concentration with ultracentrifugation through a 20% sucrose cushion. The vector titers are determined by fluorescence- activated cell sorting (FACS) analysis or by real time qPCR. The production and titration of lentiviral vectors in this protocol can be finished with 9 days. We provide an example of transducing these vectors into murine neocortical cultures containing both neurons and astrocytes. We demonstrate that lentiviral vectors support high efficiency of transduction and cell type-specific gene expression in primary cultured cells from CNS.

In this protocol we describe production, purification and titration of lentiviral vectors. We provide an example of lentiviral vector-mediated gene delivery in primary cultured neurons and astrocytes. Our methods may also apply to other cell types in vitro and in vivo.

Efficient gene delivery in the central nervous system (CNS) is important in studying gene functions, modeling neurological diseases and developing ...

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

In this article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the lipophilic fluorescent membrane marker DiI. This method allows the visualization of neuronal cell morphology in great detail. It is possible to label any cell in the CNS: cell bodies of target neurons are visualized under DIC optics or by expression of a fluorescent genetic marker such as GFP. After labeling, the DiI can be transformed into a permanent brown stain by photoconversion to allow visualization of cell morphology with transmitted light and DIC optics. Alternatively, the DiI-labeled cells can be observed directly with confocal microscopy, enabling genetically introduced fluorescent reporter proteins to be colocalised. The technique can be used in any animal, irrespective of genotype, making it possible to analyze mutant phenotypes at single cell resolution.

Labeling of Single Cells in the Central Nervous System of Drosophila melanogaster

We present a technique for labeling single neurons in the central nervous system (CNS) of Drosophila embryos, which allows the analysis of neuronal morphology by either transmitted light or confocal microscopy.

In this  article we describe how to individually label neurons in the embryonic CNS of Drosophila melanogaster by juxtacellular injection of the ...

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. Understanding repair and regeneration in animals is a key question in biology. The damaged human CNS does not regenerate, and understanding how to promote the regeneration is one of main goals of medical neuroscience. The powerful genetic toolkit of Drosophila can be used to tackle the problem of CNS regeneration.
A lesion to the CNS ventral nerve cord (VNC, equivalent to the vertebrate spinal cord) is applied manually with a tungsten needle. The VNC can subsequently be filmed in time-lapse using laser scanning confocal microscopy for up to 24 hr to follow the development of the lesion over time. Alternatively, it can be cultured, then fixed and stained using immunofluorescence to visualize neuron and glial cells with confocal microscopy. Using appropriate markers, changes in cell morphology and cell state as a result of injury can be visualized. With ImageJ and purposely developed plug-ins, quantitative and statistical analyses can be carried out to measure changes in wound size over time and the effects of injury in cell proliferation and cell death. These methods allow the analysis of large sample sizes. They can be combined with the powerful genetics of Drosophila to investigate the molecular mechanisms underlying CNS regeneration and repair.

An Injury Paradigm to Investigate Central Nervous System Repair in Drosophila

An injury paradigm using the Drosophila larval ventral nerve cord to investigate central nervous system regeneration and repair is described. Stabbing followed by laser scanning confocal microscopy in time-lapse and fixed specimens, combined with quantitative analysis with purposefully developed software and genetics, are used to investigate the molecular mechanisms of CNS regeneration and repair.

An experimental method has been developed to investigate the cellular responses to central nervous system (CNS) injury using the fruit-fly Drosophila. ...

Direct Intraventricular Delivery of Drugs to the Rodent Central Nervous System

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses specifically on antisense oligonucleotides (ASOs), though the techniques shown in the video here can also be used to deliver a plethora of other drugs to the CNS. Antisense oligonucleotides (ASOs) have the capability to knockdown sequence-specific targets 1 as well as shift isoform ratios of specific genes 2. To achieve widespread gene knockdown or splicing in the CNS of mice, the ASOs can be delivered into the brain using two separate routes of administration, both of which we demonstrate in the video.
The first uses Alzet osmotic pumps, connected to a catheter that is surgically implanted into the lateral ventricle. This allows the ASOs to be continuously infused into the CNS for a designated period of time. The second involves a single bolus injection of a high concentration of ASO into the right lateral ventricle. Both methods use the mouse cerebral ventricular system to deliver the ASO to the entire brain and spinal cord, though depending on the needs of the study, one method may be preferred over the other.

We describe a method to target drugs to the central nervous system by either implanting a catheter or performing a bolus injection into the right lateral ventricle in mice. We focus specifically on the delivery of antisense oligonucleotides. This technique is readily adaptable to other drugs and to rats.

Due to an inability to cross the blood brain barrier, certain drugs need to be directly delivered into the central nervous system (CNS). Our lab focuses ...

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution has been a long sought tool for neuroscientists in the systems-level search for the neural circuitry governing complex behavioral states. Optogenetics is a cutting-edge tool that is revolutionizing the field of neuroscience and represents one of the first systematic approaches to enable causal testing regarding the relation between neural signaling events and behavior. By combining optical and genetic approaches, neural signaling can be bi-directionally controlled through expression of light-sensitive ion channels (opsins) in mammalian cells. The current protocol describes delivery of specific wavelengths of light to opsin-expressing cells in deep brain structures of awake, freely-moving rodents for neural circuit modulation. Theoretical principles of light transmission as an experimental consideration are discussed in the context of performing in vivo optogenetic stimulation. The protocol details the design and construction of both simple and complex laser configurations and describes tethering strategies to permit simultaneous stimulation of multiple animals for high-throughput behavioral testing.

In vivo Optogenetic Stimulation of the Rodent Central Nervous System

Optogenetics has become a powerful tool for use in behavioral neuroscience experiments. This protocol offers a step-by-step guide to the design and set-up of laser systems, and provides a full protocol for carrying out multiple and simultaneous in vivo optogenetic stimulations compatible with most rodent behavioral testing paradigms.

The ability to probe defined neural circuits in awake, freely-moving animals with cell-type specificity, spatial precision, and high temporal resolution ...

Functional Evaluation of Biological Neurotoxins in Networked Cultures of Stem Cell-derived Central Nervous System Neurons

Therapeutic and mechanistic studies of the presynaptically targeted clostridial neurotoxins (CNTs) have been limited by the need for a scalable, cell-based model that produces functioning synapses and undergoes physiological responses to intoxication. Here we describe a simple and robust method to efficiently differentiate murine embryonic stem cells (ESCs) into defined lineages of synaptically active, networked neurons. Following an 8 day differentiation protocol, mouse embryonic stem cell-derived neurons (ESNs) rapidly express and compartmentalize neurotypic proteins, form neuronal morphologies and develop intrinsic electrical responses. By 18 days after differentiation (DIV 18), ESNs exhibit active glutamatergic and &#947;-aminobutyric acid (GABA)ergic synapses and emergent network behaviors characterized by an excitatory:inhibitory balance. To determine whether intoxication with CNTs functionally antagonizes synaptic neurotransmission, thereby replicating the in vivo pathophysiology that is responsible for clinical manifestations of botulism or tetanus, whole-cell patch clamp electrophysiology was used to quantify spontaneous miniature excitatory post-synaptic currents (mEPSCs) in ESNs exposed to tetanus neurotoxin (TeNT) or botulinum neurotoxin (BoNT) serotypes /A-/G. In all cases, ESNs exhibited near-complete loss of synaptic activity within 20 hr. Intoxicated neurons remained viable, as demonstrated by unchanged resting membrane potentials and intrinsic electrical responses. To further characterize the sensitivity of this approach, dose-dependent effects of intoxication on synaptic activity were measured 20 hr after addition of BoNT/A. Intoxication with 0.005 pM BoNT/A resulted in a significant decrement in mEPSCs, with a median inhibitory concentration (IC50) of 0.013 pM. Comparisons of median doses indicate that functional measurements of synaptic inhibition are faster, more specific and more sensitive than SNARE cleavage assays or the mouse lethality assay. These data validate the use of synaptically coupled, stem cell-derived neurons for the highly specific and sensitive detection of CNTs.

A custom protocol is described to differentiate mouse ES cells into defined populations of highly pure neurons exhibiting functioning synapses and emergent network behavior. Electrophysiological analysis demonstrates the loss of synaptic transmission following exposure to botulinum neurotoxin serotypes /A-/G and tetanus neurotoxin.

Therapeutic and mechanistic studies of the presynaptically targeted clostridial neurotoxins (CNTs) have been limited by the need for a scalable, ...

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise from extra-retinal sources. Ex vivo ERG provides an efficient method to dissect the function of retinal cells directly from an intact isolated retina of animals or donor eyes. In addition, ex vivo ERG can be used to test the efficacy and safety of potential therapeutic agents on retina tissue from animals or humans. We show here how commercially available in vivo ERG systems can be used to conduct ex vivo ERG recordings from isolated mouse retinas. We combine the light stimulation, electronic and heating units of a standard in vivo system with custom-designed specimen holder, gravity-controlled perfusion system and electromagnetic noise shielding to record low-noise ex vivo ERG signals simultaneously from two retinas with the acquisition software included in commercial in vivo systems. Further, we demonstrate how to use this method in combination with pharmacological treatments that remove specific ERG components in order to dissect the function of certain retinal cell types.

Simultaneous ex vivo Functional Testing of Two Retinas by in vivo Electroretinogram System

Ex vivo ERG can be used to record electrical activity of retinal cells directly from isolated intact retinas of animals or humans. We demonstrate here how common in vivo ERG systems can be adapted for ex vivo ERG recordings in order to dissect the electrical activity of retinal cells.

An In vivo electroretinogram (ERG) signal is composed of several overlapping components originating from different retinal cell types, as well as noise ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...